The Chromosomal mazEF Locus of Streptococcus mutans Encodes a Functional Type II Toxin-Antitoxin Addiction System

Abstract

Type II chromosomal toxin-antitoxin (TA) modules consist of a pair of genes that encode two components: a stable toxin and a labile antitoxin interfering with the lethal action of the toxin through protein complex formation. Bioinformatic analysis of Streptococcus mutans UA159 genome identified a pair of linked genes encoding a MazEF-like TA. Our results show that S. mutans mazEF genes form a bicistronic operon that is cotranscribed from a σ70-like promoter. Overproduction of S. mutans MazF toxin had a toxic effect on S. mutans which can be neutralized by coexpression of its cognate antitoxin, S. mutans MazE. Although mazF expression inhibited cell growth, no cell lysis of S. mutans cultures was observed under the conditions tested. The MazEF TA is also functional in E. coli, where S. mutans MazF did not kill the cells but rather caused reversible cell growth arrest. Recombinant S. mutans MazE and MazF proteins were purified and were shown to interact with each other in vivo, confirming the nature of this TA as a type II addiction system. Our data indicate that MazF is a toxic nuclease arresting cell growth through the mechanism of RNA cleavage and that MazE inhibits the RNase activity of MazF by forming a complex. Our results suggest that the MazEF TA module might represent a cell growth modulator facilitating the persistence of S. mutans under the harsh conditions of the oral cavity.

Most prokaryotic chromosomes contain a number of toxic or suicidal genes that induce reversible cell growth arrest or death (18). These chromosomal toxin-antitoxin (TA) modules consist of a pair of genes that encode two components: a protein toxin and an antitoxin (labile protein or untranslated antisense RNA) that interferes with the lethal action of the toxin (15). Typically, the toxin inhibits an essential cell function such as translation or DNA replication (18). The TA modules are classified into two major types based on the nature of the antitoxin, the toxin always being a protein. Type I TA systems consist of an RNA antitoxin and a protein toxin, in which the RNA antitoxin inhibits translation of the toxin mRNA (13, 14). In type II TA systems, both toxins and antitoxins are proteins, where the unstable antitoxin neutralizes the toxicity of the cognate toxin by forming a harmless complex (for a recent review, see reference 32). In all studied type II TA systems, the protein antitoxin has a much shorter in vivo half-life than the toxin and is degraded by a specific intracellular protease such as Lon or Clp (6). TA systems are also known as addiction modules as cells become “addicted” to the short-lived antitoxin product (the unstable antitoxin is degraded faster than the more stable toxin) since its de novo synthesis is essential for their survival (9). All reported type II TA operons are autoregulated at the level of transcription by the antitoxin or the concerted action of toxin and antitoxin binding to their promoter (32). The type II toxins characterized thus far include protein kinases, phosphotransferases, DNA gyrase poisons, and ribosome-dependent and ribosome-independent ribonucleases (45). The vast majority of type II toxins are mRNA-specific endonucleases, also called mRNA interferases. Importantly, chromosomal type II TAs induce a reversible bacteriostatic state since no toxin has been reported to be directly bacteriolytic (7, 32, 38, 43).

TA modules are ubiquitous in the chromosomes of archaea and bacteria, often found in multiple copies (32, 37). Because TA systems are thought to invade prokaryotic genomes through horizontal gene transfer and intragenomic recombination, they are commonly described as mobile genetic elements (32). Comprehensive genome analyses have revealed the diversity in the distribution of TA systems. Among bacteria, Bacteroidetes/Chlorobi, Alphaproteobacteria, Gammaproteobacteria, and Cyanobacteria possess the greatest variety of TAs, while Firmicutes are characterized by a particularly low TA diversity (32, 37). Recently, Makarova et al. (32) reported the most detailed and comprehensive comparative analysis of type II TAs in 750 complete genomes of archaea and bacteria. This analysis resulted in the prediction of 12 new families of putative toxins and 13 new families of putative antitoxins. Since these small modules are highly diverse and broadly distributed in prokaryotes, one can postulate that they might have multiple biological roles. For example, the fact that almost all obligate intracellular microorganisms (endosymbiotic species) are devoid of TA loci, whereas free-living prokaryotes have several TA loci, has lead some researchers to propose the hypothesis that TAs are stress-response elements that help free-living prokaryotes to cope with environmental stress (6, 15). Indeed, obligate intracellular microorganisms thrive in constant environments and are thus expected to encounter minimal stress or environmental fluctuations. Nevertheless, the physiological role of chromosomally encoded TAs still remains the subject of debate. Different hypotheses have been proposed to explain the role of TAs in bacterial physiology, including stress survival, growth control, programmed cell death, persister formation, stabilization of genome, and anti-addiction system (7, 8, 28, 31, 43, 45).

Streptococcus mutans is a member of the division Firmicutes. In the presence of fermentable dietary carbohydrates, this acid-producing bacterium present in the oral biofilm can cause damage (cavities) to the tooth's hard tissues (4, 12, 34). Although tooth decay is largely preventable, it remains the most common and costly disease worldwide. By sequence homology, at least four type II TA systems have been identified in the S. mutans UA159 reference strain (3, 32, 40). In an attempt to investigate the physiological roles of S. mutans TA modules, mutants lacking MazF (SMU.173), RelE (SMU.896), or both putative toxins were constructed and tested for their impact on the capacity of S. mutans to form biofilm and withstand acid killing (26). Interestingly, when both putative toxin-encoding genes were inactivated, acid tolerance and growth properties of S. mutans were affected. These results led Burne and coworkers (26) to propose that chromosomal TA modules of S. mutans could function as regulators of cell growth in response to environmental stresses. We provide here definitive evidence that the SMU.172-SMU.173 locus of S. mutans encodes a functional type II TA addiction system, the first characterized TA module in an oral pathogen.

MATERIALS AND METHODS

Bacterial strains and growth conditions.

A summary of bacterial strains and plasmids used in the present study is provided in Table 1 . S. mutans UA159 wild-type strain and its mutants were grown in Todd-Hewitt yeast extract (THYE) broth and incubated statically at 37°C in air with 5% CO₂. Mutants were constructed in UA159 wild-type as described previously (25). E. coli strains were grown aerobically in Luria-Bertani (LB) medium at 37°C. When needed, antibiotics were added as follows: ampicillin (100 μg/ml), chloramphenicol (20 μg/ml), or kanamycin (50 μg/ml) for E. coli and chloramphenicol (10 μg/ml), kanamycin (300 μg/ml), erythromycin (10 μg/ml), or spectinomycin (1000 μg/ml) for S. mutans. Cell growth was monitored based on the optical density at 600 nm (OD₆₀₀). Cell viability was assessed by counting CFU on replica agar plates.

TABLE 1.

Bacterial strains and plasmids used in the study

Strain or plasmid	Relevant characteristic(s)a	Source or reference
Strains
S. mutans
UA159	Wild-type reference strain	Lab stock
ΔmazEF mutant	In-frame mazEF deletion mutants derived from UA159; Emr or Spr	This study
ΔmazF mutant	In-frame mazF deletion mutants derived from UA159; Emr or Spr	This study
UA159(pIB187)	UA159 harboring pIB187; Emr	This study
ΔmazEF(pIB187) mutant	ΔmazEF mutant harboring pIB187; Spr Emr	This study
E. coli
DH10B	Host strain for cloning and plasmid production	Lab stock
TOP10	Host strain for cloning and plasmid production	Invitrogen
LMG194	Host strain for pBAD expression	Invitrogen
BL21(DE3)	Host strain for pET15TV-L expression	Novagen
DH10B(pMAS1)	DH10B containing pMAS1; Kmr	This study
DH10B(pMAS1/pIB166)	DH10B containing pMAS1/pIB166; Kmr Cmr	This study
DH10B(pMAS1/pMAS2)	DH10B containing pMAS1/pMAS2; Kmr Cmr	This study
DH10B(pMAS1/pMAS3)	DH10B containing pMAS1/pMAS3; Kmr Cmr	This study
TOP10(pMAS4)	TOP10 containing pMAS4; Kmr	This study
TOP10(pMAS5)	TOP10 containing pMAS5; Kmr	This study
LMG194(pBAD202/D/lacZ)	LMG194 containing pBAD202/D/lacZ; Kmr	This study
LMG194(pMAS4)	LMG194 containing pMAS4; Kmr	This study
LMG194(pMAS5)	LMG194 containing pMAS5; Kmr	This study
BL21(DE3)(pMAS4)	BL21(DE3) containing pMAS4; Kmr	This study
BL21(DE3)(pMAS6)	BL21(DE3) containing pMAS6; Apr	This study
BL21(DE3)(pMAS4/pMAS6)	BL21(DE3) containing pMAS4/pMAS6; Kmr Apr	This study
Plasmids
pIB166	Shuttle plasmid containing the P23 lactococcal promoter; Cmr	5
pIB187	Shuttle plasmid containing the gusA gene encoding the β-glucuronidase enzyme under the control of P23; Emr	5
pHSG299	High-copy-number cloning vector; Kmr	Takara Bio USA
pBAD202/D-TOPO	Expression vector linearized and topoisomerase-activated; Kmr	Invitrogen
pBAD202/D/lacZ	Control plasmid encoding the β-galactosidase LacZ	Invitrogen
pET15TV-L	Expression vector derived from pET15b; Apr	Lab stock
pMAS1	mazE cloned under the control of the IPTG inducible lac promoter into pHSG299; Kmr	This study
pMAS2	mazEF cloned into pIB166; Cmr	This study
pMAS3	mazF gene cloned into pIB166; Cmr	This study
pMAS4	mazF cloned under the control of araBAD promoter into pBAD202/D-TOPO vector; Kmr	This study
pMAS5	mazEF cloned under the control of araBAD promoter into pBAD202/D-TOPO vector; Kmr	This study
pMAS6	mazE cloned under the control of T7 promoter into pET15TV-L vector; Apr	This study

^aAp^r, ampicillin resistance; Em^r, erythromycin resistance; Cm^r, chloramphenicol resistance; Km^r, kanamycin resistance; Sp^r, spectinomycin resistance.

RT-PCR.

To confirm the genetic organization and transcriptional coupling between S. mutans mazE and mazF genes, reverse transcription-PCR (RT-PCR) was performed with the primers CMT-264 (5′-ACTGTAGAAGATCTGTTTAAAG-3′) and CMT-265 (5′-AAAATCGATTGTAACCAGTTGG-3′) spanning the 3′ end and 5′ end of antitoxin- and toxin-encoding genes, respectively. Total RNA was isolated from mid-log-phase UA159 cultures (OD₆₀₀ ∼ 0.4) using TRIzol reagent (Invitrogen), DNase treated, and converted to cDNA by using a First-Strand cDNA synthesis kit (MBI Fermentas). PCR was performed with 800 ng of cDNA. Controls without reverse transcriptase enzyme were included in all experiments. As a positive control, the same RT-PCR primers were used to directly amplify the genomic DNA (gDNA) of UA159. PCR products of 323 bp were resolved on a 1.0% (wt/vol) agarose gel.

5′RACE-PCR.

A 5′ rapid amplification of cDNA ends (5′RACE)-PCR technique was used to define the transcriptional start site of the mazEF operon. Total RNA was isolated from mid-log-phase UA159 cultures by using TRIzol reagent (Invitrogen). DNA-free RNA (10 μg) was reverse transcribed by using the RACE outer primer (5′-TTACCATTGTTCATTTCCCACA-3′) and Superscript II reverse transcriptase (Invitrogen) according to the supplier's instructions. RNase H and RNase T₁ (Ambion) were then added, followed by incubation at 37°C for 30 min. The cDNA was purified by using a StrataPrep PCR purification kit (Stratagene) according to the manufacturer's instructions. Tailing of purified cDNA using the terminal deoxynucleotidyltransferase (Sigma) and dGTP/dTTP was performed according to the manufacturer's instructions. Tailed cDNAs were amplified by PCR using RACE universal primers (5′-GAATTCGAATTCCCCCCCCCCCC-3′ and 5′-GAATTCGAATTCAAAAAAAAAAAA-3′) and a RACE inner primer (5′-CTTTAAACAGATCTTCTACAGT-3′). The amplicons were analyzed by agarose gel electrophoresis and sequenced using the RACE inner primer.

Expression vectors for induction in S. mutans.

The full-length coding region of the S. mutans mazEF and mazF genes were first PCR amplified using UA159 gDNA as a template and the primers CMT-392 (5′-GAATTCGAAATAGGAGGTGGCATTATG-3′) and CMT-388 (5′-AAGCTTAGATTTCAGTAGAAGTTTAAC-3′) for the mazEF operon and the primers CMT-387 (5′-GAATTCAAGTGTGGGAAATGAACAATG-3′) and CMT-388 for the mazF gene. The purified PCR products were double digested with EcoRI/HindIII and cloned under the control of the lactococcal promoter P₂₃ in the shuttle plasmid pIB166 (5) precut by the same enzymes. The recombinant plasmids pMAS2 (mazEF in pIB166) and pMAS3 (mazF in pIB166) were transferred into electrocompetent DH10B and DH10B(pMAS1) cells, respectively. The recombinant plasmids were sequenced on both strands for verification. The plasmids pIB166, pMAS2, and pMAS3 were then used for the toxicity assays.

Toxicity assays: S. mutans transformation.

S. mutans UA159 overnight cells were diluted (1:20) with fresh THYE, grown to an OD₆₀₀ of ∼0.1, and divided into three 0.5-ml aliquots of (i) pIB166 (control), (ii) pMAS2 (mazEF in pIB166), and (iii) pMAS3 (mazF in pIB166). Portions (10 μg) of plasmids were added to the cultures, which were grown for a further 2.5 h at 37°C with 5% CO₂ in the presence of S. mutans synthetic competence-stimulating peptide at a final concentration of 0.5 μg/ml. Cultures were serially diluted and plated on THYE agar. The transformation efficiency (TE) was calculated as the percentage of chloramphenicol-resistant transformants divided by the total number of recipient cells, which was determined by the number of CFU on antibiotic-free THYE agar plates. All assays were performed in triplicate from three independent experiments. Statistical significance was determined by using a Student t test and a P value of <0.01.

Cell lysis.

The β-glucuronidase (GUS) enzyme was chosen as the marker enzyme for the cell lysis assay since it is known to be rather stable against proteolytic degradation and there has been no indication that the expression of this protein is toxic (35). Cell lysis was assessed by harvesting the supernatant of cultures expressing the gusA reporter gene cloned into pIB187 as described previously (39). Briefly, supernatants were combined in equal parts with 2× GUS buffer (100 mM Na₂HPO₄ [pH 7.0], 20 mM β-mercaptoethanol, 2 mM EDTA, 0.2% Triton X-100, 2 mM para-nitrophenyl-β-d-glucuronide [PNPG] substrate; Sigma). The reaction was incubated at 37°C for 180 min and stopped by the addition of 100 mM Na₂CO₃. The absorbance at 420 nm (A₄₂₀) was measured after 180 min of color development. The GUS activity was expressed as (1,000 × A₄₂₀)/(time in min × OD₆₀₀) in Miller units (MU).

Production and purification of recombinant fusion proteins in E. coli.

The full-length coding region of the S. mutans mazF gene and mazEF operon were first PCR amplified using UA159 gDNA as a template and the primers CMT-445 (5′-CACCATGGTAACCATCAAGCAAGG-3′) and CMT-451 (5′-TCCTTTTTCAAATAGCACCTTG-3′) for the mazF gene and CMT-444 (5′-CACCATGCAACTAGTCATCAATAAAT-3′) and CMT-451 for the mazEF operon. The PCR products were purified, cloned in-frame upstream from the His₆ sequence into pBAD202/D-TOPO vector (Invitrogen), and transferred into TOP10 chemically competent E. coli cells. The recombinant plasmids were sequenced on both strands for confirmation. The plasmids designated pMAS4 and pMAS5 were then transferred into electrocompetent E. coli LMG194 (Invitrogen) cells to express the His₆-tagged MazF (rMazF-His₆) and His₆-tagged MazEF (rMazEF-His₆) recombinant proteins, respectively. Overnight, LMG194(pMAS4) and LMG194(pMAS5) cells were diluted (1:100) into fresh LB medium supplemented with kanamycin at 50 μg/ml and grown aerobically at 37°C until an OD₆₀₀ of ∼0.5 was reached. Arabinose was then added at a final concentration of 0.02% to induce the expression of recombinant fusion proteins and the incubation was continued for another 4 h at 37°C with agitation.

The cells were collected by centrifugation, resuspended in 1× binding buffer (Novagen), and disrupted on ice by sonication. The soluble fractions of the disrupted cells were recovered by centrifugation, and the rMazF-His₆ and rMazEF-His₆ proteins were then purified by affinity chromatography on Ni²⁺-nitrilotriacetic acid (Ni-NTA) resin (Novagen) as described by the manufacturer. The Bradford protein assay was used to determine the protein concentration of the purified samples.

Protein electrophoresis.

SDS-PAGE was performed using the buffer system of Laemmli at a constant voltage (200 V) with gels containing 12% polyacrylamide in the separating gel and 4.5% polyacrylamide in the stacking gel. The protein bands were visualized by staining with Coomassie brilliant blue. For native PAGE, the Laemmli's gel system was used without SDS and β-mercaptoethanol. The proteins were transferred to a nitrocellulose membrane as described previously (27), and the recombinant proteins were detected by Western blotting with an anti-His₆ monoclonal antibody (Sigma).

In-gel digestion and mass spectrometry.

Matrix-assisted laser desorption ionization-time of flight mass spectrometry was performed at the Proteomics Platform, Quebec Genomics Center (Québec City, Québec, Canada). Briefly, protein bands of interest were extracted from the gels, placed in 96-well plates, and then washed with water. Tryptic digestion was performed on a MassPrep liquid handling robot (Waters, Milford, CT) according to the manufacturer's specifications and the protocol of Shevchenko et al. (41) with the modifications suggested by Havlis et al. (17). Peptide samples were separated by using online reversed-phase nanoscale capillary liquid chromatography and analyzed by electrospray-tandem mass spectrometry. Database searches were conducted by using Mascot software (version 2.2.0; Matrix Science, London, United Kingdom) on an S. mutans-specific database containing protein sequences deduced from the genome of UA159 reference strain (GenBank accession number AE014133).

RNase activity.

Total RNA was extracted from S. mutans UA159 as described above. Total RNA was extracted from E. coli LMG194 cultures (OD₆₀₀ ∼ 0.5) using the RNeasy Minikit (Qiagen), and isolated RNA preparations were treated with RQ1 RNase-Free DNase (Promega) according to the manufacturer's protocol. DNase-treated RNA substrates were incubated with rMazF at 37°C for 15 min. The purified recombinant β-galactosidase (rLacZ) protein was used in the control experiments. Each reaction mixture contained 4 μg of DNase-treated RNA substrate, 20 pmol of rMazF, and 1× nuclease buffer (20 mM Tris-HCl [pH 8], 100 mM NaCl, 1 mM dithiothreitol). The reaction mixture was then subjected to 1.2% agarose gel electrophoresis. Inhibition of cleavage with the addition of rMazE antitoxin was also tested. Briefly, 4 μg of DNase-treated RNA substrate was incubated in 1× nuclease buffer at 37°C for 15 min with rMazE and rMazF in different ratio combinations.

RESULTS

Sequence analysis of the chromosomal mazEF locus.

DNA sequence analysis of the S. mutans UA159 genome revealed that the SMU.172-SMU.173 locus encodes a putative TA pair (3). BLASTP analysis revealed that SMU.172 encodes a putative protein of 81 amino acids. PSI-BLAST search revealed a statistically significant similarity (E-value, 7.9e⁻⁰⁶) with the AbrB/MazE superfamily of DNA-binding proteins. AbrB proteins have been identified as antitoxins in well-characterized type II TA systems such as MazEF, Kis-Kid, and PemIK (32). Apparently arranged in a bicistronic operon, the SMU.173 open reading frame overlaps with SMU.172 by seven nucleotides (Fig. 1A). Bioinformatic analysis of SMU.173 revealed that the putative protein of 110 amino acids belongs to the PemK/MazF interferase superfamily of toxins (E-value, 4.84e⁻¹²). This family consists of growth inhibitor proteins such as PemK, ChpA, ChpB, Kid, and MazF (32). Thus, the S. mutans SMU.172-SMU.173 locus was investigated to determine whether the two genes encoded for a functional type II TA system.

FIG. 1.

(A) Nucleotide and deduced amino acid sequences of the mazEF (SMU.172-SMU.173) locus. Putative −35 and −10 promoter sites and putative ribosome-binding sites (RBS) are indicated. A factor-independent terminator-like structure is underlined with broken lines. The transcriptional start site (+1) identified by 5′RACE-PCR is indicated below the sequence. (B) Agarose gel electrophoresis of the RT-PCR amplification product. Lanes: L, 1-kb Plus DNA Ladder (Invitrogen); 1, positive control (UA159 gDNA); 2, negative control (no RT); 3, RT-PCR (UA159 cDNA).

To confirm the genetic organization and transcriptional coupling between mazE (SMU.172) and mazF (SMU.173) genes, RT-PCR was performed using primers spanning the 3′ end and 5′ end of antitoxin- and toxin-encoding genes, respectively. A single PCR product of 323 bp was amplified and sequenced, confirming that the mazEF mRNA was bicistronic (Fig. 1B). Using 5′RACE-PCR, we determined the transcriptional start site of the mazEF operon to be located seven nucleotides 3′ proximal to the inferred Pribnow box (TATAAT), suggesting that mazEF comprises a transcriptionally discrete locus in the S. mutans UA159 genome.

The mazEF locus encodes a functional TA module.

The lack of a tightly regulated inducible expression system for S. mutans prompted us to exploit the ability of S. mutans to become naturally competent to design our toxicity assay. Naturally competent bacteria have the ability to take free DNA from the surrounding medium and incorporate this DNA into their genomes by homologous recombination, a phenomenon first discovered in Streptococcus pneumoniae (16). S. mutans strains behave like pneumococci since they are naturally competent for transformation (29). The toxicity of the predicted S. mutans MazF toxin in its native host was tested by natural transformation of the ΔmazEF and ΔmazF mutants (UA159 background) with the empty plasmid (pIB166), the plasmid pIB166 containing mazEF operon driven by the constitutive P₂₃ promoter (pMAS2), and the plasmid pIB166 containing mazF gene under the control of P₂₃ (pMAS3). It is worth mentioning that despite several attempts, the mazF gene could be cloned into pIB166 plasmid and expressed in E. coli background only when the MazE antitoxin was preliminarily induced. Consequently, the mazE gene was cloned under the control of an IPTG (isopropyl-β-d-thiogalactopyranoside)-inducible promoter into the pHGS299 vector (a plasmid unable to replicate in streptococci). The E. coli cells were made electrocompetent and used as recipient cells to propagate pMAS3 plasmid necessary for the toxicity assay. The transformation efficiencies of the ΔmazEF mutant were (7.2 ± 1.0) × 10⁻² and (8.0 ± 0.3) × 10⁻² for pIB166 (empty vector) and pMAS2 (MazEF), respectively, whereas no colonies were recovered when ΔmazEF cells were transformed with pMAS3 expressing the MazF toxin alone (Table 2). In contrast, the ΔmazF mutant benefited from the presence of the wild-type mazE copy into its genome to protect the cells since plasmid pMAS3 was able to replicate. Indeed, the transformation efficiency of the ΔmazF mutant for pMAS3 was similar to those obtained with the pIB166 vector and the pMAS2 construct (Table 2). These results clearly showed a growth-inhibitory effect of MazF toxin on S. mutans cells and that the toxic effect elicited by MazF can be neutralized by coexpression of its cognate MazE antitoxin. Hence, mazEF operon encodes a functional TA system in S. mutans.

TABLE 2.

S. mutans toxicity assay based on natural competence

Construct	Description	TE ± SDa
		ΔmazEF mutant	ΔmazF mutant
pIB166	Empty vector	(7.2 ± 1.0) × 10−2	(12.2 ± 3.1) × 10−2
pMAS2	mazEF genes	(8.0 ± 0.3) × 10−2	(11.1 ± 3.7) × 10−2
pMAS3	mazF gene	0b	(7.5 ± 1.0) × 10−2

^aThe transformation efficiency (TE) was calculated as the percentage of transformants over the total number of recipient cells.

^bP < 0.01, as determined using a Student t test comparing results for the ΔmazEF mutant for pMAS3 to those for the ΔmazEF mutant for pIB166.

Effect of MazF mutations on toxicity.

As previously mentioned, all our attempts at cloning the S. mutans mazF gene without its cognate antitoxin gene in E. coli gave rise to mutated S. mutans mazF clones, and most of these amino acid changes occurred at conserved positions (see Fig. S1 in the supplemental material). In order to test the importance of these amino acid mutations in the functionality of the MazF protein, we tested randomly selected clones (Table 3) in our toxicity assay (natural transformation). Dramatic effects were observed with the substitutions I29F, L61Q, E21K, and F45L since all four point mutations restored the transformation efficiency to the wild-type level (Fig. 2). Although clones S193, S195, and S197 did not restore the transformation efficiency to full wild-type levels, the decrease in transforming ability was not considered statistically significant. We can hypothesize that these point mutations substantially reduced or completely abolished the toxic activity of MazF toxin since cells of the naturally transformable ΔmazEF mutant were able to replicate the MazF recombinant plasmids.

TABLE 3.

Amino acid modifications of randomly selected MazF clones

Clone	Amino acid modification(s)
S193	R53H, R102W
S194	I29F
S195	K11E
S196	L61Q
S197	Q74R
S198	E21K
S199	F45L
S200	N41I, Q74L

FIG. 2.

Effect of MazF mutations on S. mutans toxicity based on natural competence. Portions (10 μg) of the plasmids were added to S. mutans ΔmazEF cultures grown to early log phase. After 2.5 h at 37°C, the cultures were serially diluted and plated. The transformation efficiency was expressed as the percentage of chloramphenicol-resistant transformants divided by the total number of recipient cells. Although clones S193, S195, and S197 did not restore transformation efficiency to full wild-type levels (C, control), the decrease in transforming ability was not considered statistically significant. All assays were performed in triplicate from three independent experiments and statistical significance was determined by using a Student t test and a P value of <0.01.

MazF toxicity does not result in S. mutans autolysis during natural transformation.

In order to investigate whether the growth-inhibitory effect of MazF toxin during natural transformation leads to lysis of S. mutans, we used the β-glucuronidase (GUS) enzyme as the marker enzyme; the release of the cytoplasmic GUS enzyme into the supernatant of transformed cells was quantified as a measure of cell lysis. The ΔmazEF mutant was thus transformed with pIB187 plasmid harboring a GUS reporter gene under the control of the P₂₃ lactococcal promoter for constitutive expression. The new S. mutans ΔmazEF(pIB187) background strain was then used in our toxicity assay (natural transformation) as described previously. As expected, the transformation efficiency was similar for pIB166 (empty plasmid) and pMAS2 (expressing both the antitoxin and the toxin genes), whereas no transformant could be detected with the pMAS3 construct containing the mazF toxin gene alone (Table 4). Interestingly, GUS activity was not significantly altered in the supernatants of cultures tested with pMAS3, suggesting that MazF toxic activity does not lead to cell lysis of S. mutans under the experimental conditions tested. Moreover, the fact that the transformation assays with pMAS3 construct did not alter the total number of recipient cells, since MazF expression had no effect on the proportion of dying cells, further supports our conclusion.

TABLE 4.

Effect of MazF on S. mutans lysis during the transformation experiments^a

Construct	TE ± SDb	GUS activityc (MU ± SD)
pIB166 (empty vector)	(1.2 ± 0.2) × 10−2	8.02 ± 0.57
pMAS2 (mazEF)	(1.8 ± 0.7) × 10−2	7.37 ± 0.77
pMAS3 (mazF)	0d	7.23 ± 0.92

^aAll assays were performed in triplicate from three independent experiments.

^bTE, transformation efficiency.

^cGUS activity was expressed in Miller units (MU). See Materials and Methods for details.

^dP < 0.01, as determined using a Student t test comparing results of the ΔmazEF(pIB187) strain for pMAS3 to those of the ΔmazEF(pIB187) strain for pIB166.

Expression of S. mutans MazF toxin leads to E. coli cell growth arrest and inhibits colony formation.

E. coli cells have been successfully used previously as a host for verification of the function of various heterologically expressed TA loci (1, 11, 21, 36, 46). The S. mutans mazF gene was thus cloned into the pBAD expression system for induction of gene expression using araBAD promoter dose-dependent regulation. The recombinant plasmid designated pMAS4 harboring the streptococcal toxin gene was used to transform E. coli LMG194, and transformants were selected on LB-kanamycin agar plates supplemented with glucose to repress the basal expression level. The growth of LMG194(pMAS4) transformants in liquid medium supplemented with 0.02% arabinose as an inducer showed significant growth differences (Fig. 3A). Strain LMG194(pMAS5), in which both MazE and MazF proteins were expressed, behaves similarly to the LMG194(pBAD202/D/lacZ) control strain. In cell viability assays, overexpression of MazF resulted in growth inhibition with a reduction of ∼10² in cell viability by 4 h postinduction compared to LMG194(pMAS5) and LMG194(pBAD202/D/lacZ) (Fig. 3B). When tested on solid media, LMG194 cells harboring pMAS4 could not grow in the presence of arabinose at 0.2%, whereas LMG194(pBAD202/D/lacZ) and LMG194(pMAS5) were able to grow (data not shown). The lack of colony formation indicates either that MazF kills the cells or that MazF induces a state wherein most cells remained viable but in which colony formation is inhibited. Nevertheless, these results demonstrated that MazF is also toxic to E. coli cells, and this toxicity is abolished by coexpression of MazE antitoxin.

FIG. 3.

Characterization of the MazEF TA in E. coli. (A) The growth characteristics of LMG194 cells expressing LacZ (pBAD202/D/lacZ), MazF (pMAS4), and MazEF (pMAS5) were analyzed by measuring the absorbance (OD₆₀₀) after every hour. (B) After arabinose induction, appropriate dilutions were plated on LB agar for determination of the number of CFU per ml.

Recovery of E. coli cell growth arrest after the induction of S. mutans MazE antitoxin in cultures treated with S. mutans MazF toxin.

To determine whether MazE antitoxin can rescue MazF-induced cell growth arrest, we tested the effect of MazE on E. coli cells in which MazF was previously overexpressed. The mazE gene was cloned into pET15TV-L under the IPTG-inducible T7 promoter. The recombinant plasmid pMAS6 was transferred into BL21(DE3) containing plasmid pMAS4 for MazF expression in the presence of arabinose. Cells of BL21(DE3) cotransformed with plasmids pMAS4 and pMAS6 were grown to mid-log phase, at which time arabinose was added to induce MazF expression. At time 2, 3, and 4 h after arabinose induction, IPTG was added to induce MazE expression for 1 h; the cells were then spread onto LB plates containing glucose to repress the pBAD promoter. As seen in Fig. 4, overexpression of MazF toxin led to a rapid decrease of CFU counts. When IPTG was added to the culture medium to induce MazE, the CFU counts increased (Fig. 4), indicating that the expression of MazE antitoxin reversed the toxicity of MazF even when the mazE reading frame was provided in trans. These results demonstrated that MazF toxin did not kill the cells but rather caused reversible cell growth arrest.

FIG. 4.

Recovery of MazF-induced cell growth arrest by MazE antitoxin. Cells of BL21DE3 cotransformed with pBAD-MazF (pMAS4) and pET15-MazE (pMAS6) were grown to mid-log phase, at which time 0.02% arabinose was added (MazF induced). At indicated time points (after arabinose induction), 1 mM IPTG was added for 1 h (MazE coinduced), after which the cells were spread on LB plates containing 0.2% glucose, 50 μg of kanamycin/ml, and 100 μg of ampicillin/ml.

MazE and MazF form a complex.

To show direct interaction between MazE antitoxin and MazF toxin, we used plasmid pMAS5, in which both MazE and MazF proteins were expressed and which was constructed so that only MazF protein was expressed as a His₆-tagged fusion protein. Recombinant proteins were expressed in LMG194 and purified by affinity chromatography. When the mixture was subjected to native PAGE, a single band was observed and detected by Western blotting with an anti-His₆ monoclonal antibody (Fig. 5A). The protein band was cut out and analyzed by mass spectrometry to identify the protein components. The results showed that the complex was composed of MazE and MazF proteins, and the molar ratio of MazE to MazF was 1:2.

FIG. 5.

Copurification and interaction of rMazE antitoxin and rMazF-His₆ toxin. (A) Native PAGE and Western blot analysis with an anti-His₆ monoclonal antibody. The protein composition of the immunoreactive band was identified by MS. Lanes: 1, LMG194(pMAS5) uninduced culture; 2, LMG194(pMAS5) arabinose-induced culture. (B) SDS-12% PAGE was performed to analyze the protein composition of the immunoreactive band. Lanes: L, PageRuler prestained protein ladder (Fermentas); 1, total cell lysate from LMG194(pMAS5) uninduced culture; 2, total cell lysate from LMG194(pMAS5) arabinose-induced culture; 3, elution of absorbed lysate from LMG194(pMAS5) uninduced culture; 4, elution of absorbed lysate from LMG194(pMAS5) arabinose-induced culture. The identity of the two distinct protein bands was confirmed by mass spectrometry. Both rMazE and rMazF-His₆ fusion proteins had the expected molecular mass: ∼ 23 kDa for rMazE, including the His-patch thioredoxin on the N-terminal end, and ∼16 kDa for rMazF-His₆, including the polyhistidine region on the C-terminal end.

When the mixture was subjected to SDS-PAGE, two distinct bands were observed that corresponded to the expected mobilities of rMazE (22.8 kDa) and rMazF-His₆ (16 kDa), thus confirming that under native conditions both rMazE and rMazF proteins were copurified (Fig. 5B). Mass spectrometry also demonstrated that both rMazE and rMazF proteins had the expected molecular mass.

MazF has RNase activity.

The PemK/MazF toxin superfamily is composed of endoribonucleases that interfere with protein synthesis by cleaving cellular mRNA. To determine whether MazF toxin has RNase activity, DNase-treated total RNA isolated from S. mutans UA159 was incubated with rMazF in a dose-dependent manner. As shown in Fig. 6, S. mutans RNA was cleaved into small fragments with 20 pmol of rMazF. The β-galactosidase fusion protein was also purified by affinity chromatography and used in a nuclease assay to confirm that the observed RNase activity was due to rMazF alone and not to contaminating E. coli RNases (Fig. 6). The RNase activity of rMazF was inhibited when the molar ratio of rMazE to rMazF was increased to 1:2, supporting the mass spectrometry results that the molar ratio of MazE-MazF complex is approximately 1:2. Similar results were obtained when the DNase-treated RNA from E. coli was used as a substrate (data not shown). These results indicated that MazF is a toxic RNase causing bacterial cell growth arrest through the mechanism of RNA cleavage and that MazE antitoxin most probably inhibits the RNase activity by forming a stable MazE-MazF complex.

FIG. 6.

RNase activity of MazF. Each reaction was performed for 15 min at 37°C with 4 μg of S. mutans DNase-treated RNA. Lanes: L, GeneRuler 1-kb Plus DNA ladder (Fermentas); No P, no protein, RNA alone; LacZ, RNA incubated with rLacZ (20 pmol); MazE, RNA incubated with rMazE (20 pmol); MazF, RNA incubated with rMazF (20 pmol); 2E:1F, RNA incubated with rMazE (40 pmol) and rMazF (20 pmol); 1E:1F, RNA incubated with rMazE (20 pmol) and rMazF (20 pmol); 1E:2F, RNA incubated with rMazE (20 pmol) and rMazF (40 pmol); 1E:4F, RNA incubated with rMazE (20 pmol) and rMazF (80 pmol).

DISCUSSION

Identification of the signaling mechanisms that regulate bacterial cell growth is critical to understanding how microorganisms colonize and persist in specific niches. The possible role of chromosomally encoded TAs in growth control is relevant to a number of important aspects of bacterial physiology, including antibiotic tolerance, maintenance of pathogen reservoirs, and chronic infections. In the present study, we reported a functional chromosomal type II TA, MazEF, in the pathogenic organism S. mutans. The mazEF locus possesses the characteristic organization of type II TAs in which the gene for the antitoxin component precedes the toxin gene and where the two genes share one or several nucleotides; overlaps have been demonstrated to be potentially important in transcriptional and translational regulators of gene expression and to influence gene evolution (20). Although most type II TAs conform to this arrangement, there are examples in which the gene order is reversed or where the product of a third gene is involved (32, 45). A number of orphan toxin genes lacking the adjacent antitoxin gene have also been found in searches for type II loci; it is possible that the synthesis of these toxins is repressed by antisense small RNAs (9, 30). For instance, a total of 45 putative toxins and antitoxins have been identified in the genome of UA159 on the basis of protein sequence and comparative genomic analysis (32), and yet only four TA pairs (AbrB/MazF, Xre/COG2856, RHH/RelE, and PIN/AbrB) possessed all of the features and could be predicted to function as bona fide type II TAs (32).

We focused our study on the functional characterization of the MazE(AbrB)/MazF system. Our results confirmed that the chromosomal mazF homologue in S. mutans encoded a toxic protein with the adjacent mazE gene encoding its cognate protein antitoxin. In S. mutans as well as in most cases, the type II toxin is a RNase that cleaves mRNA, leading to the inhibition of protein synthesis and cell growth arrest. We therefore propose that the S. mutans chromosomal MazEF module represents a growth modulator system that induces under particular stress conditions a reversible dormancy state to enhance fitness and competitiveness. Indeed, the oral cavity is a complex ecological system in which bacteria must overcome a wide range of conditions in order to survive (22, 33, 42). Previous work done by Lemos et al. (26) showed that the deletion of both mazEF and relBE in S. mutans increased resistance to low pHs when the cells were grown in biofilm. Although the molecular mechanisms underlying this process are still not known, these data are consistent with the growth modulator model. Recently, preliminary work done in our lab suggested that the stochastic expression of the MazF toxin in a small fraction of S. mutans population could lead to dormancy, allowing dormant cells to survive lethal stresses, including exposure to bactericidal antibiotics. These preliminary data are in accordance with the present study demonstrating that MazF does not cause cell lysis but inhibits cell growth by cleaving single-stranded RNA. By stopping bacterial growth, MazF could allow stressed cells to remain in a dormant or nongrowing stress-tolerant state until more favorable environmental conditions return.

One group presented evidence that MazEF of E. coli was involved in bacterial programmed cell death, a form of apoptosis (10). Indeed, Engelberg-Kulka and coworkers demonstrated that in E. coli, MazEF is a stress-induced suicidal TA module (19), and yet MazEF-induced PCD is still subject to debate since it has not been reproduced by other groups (44). Recently, Kolodkin-Gal et al. (23, 24) demonstrated that E. coli MazF-induced death was dependent on the bacterial cell density and required an extracellular death factor (EDF) in order for the stresses to be effective. The role of the EDF peptide resembles to some extent the function of the peptide pheromone CSP involved in S. mutans quorum sensing. Our lab previously demonstrated that CSP was involved in stress response and induced a dose-dependent growth inhibition of S. mutans cultures. Interestingly, cell lysis could be observed in a small fraction of the population at high concentrations of CSP (39). We sought to determine whether S. mutans could integrate its stress response with the MazEF TA via the CSP pheromone. To test the impact of MazEF on the stress response directly, ΔmazEF mutant cells were exposed to a protein synthesis-inhibitor antibiotic and acid stress (conditions known to induce expression of CSP). When exposed to sub-MICs of spectinomycin or to pH 5.0, ΔmazEF cells behave like the wild type since no significant increase or decrease in the lag phase before recovering from the stress was observed between the two strains. Although these results could not allow us to make an intimate connection between the CSP-induced stress regulon and MazEF, we cannot rule out the contribution of CSP in MazF-induced stasis under different stresses. Further experiments will be required to explore this possibility.

Assuming that chromosomal TAs are functioning as regulators of bacterial growth in response to environmental stress, we can speculate that TAs could limit bacterial growth upon encountering eukaryotic host cells (hostile environment) and allow pathogens to persist into specific niches. As a result, pathogens can persist several months (or sometimes years) and give rise to recurrent infections exceedingly difficult to eradicate. Consequently, the understanding of bacterial growth regulation with particular emphasis on the mechanism of action of TA-induced growth arrest represents a promising avenue for the development of novel and effective antimicrobial strategies.